To evaluate the in vitro antimicrobial activity of aqueous and methanol extracts of Odina wodier bark (OWB), a folk medicine, against representative bacteria, fungi and herpes simplex virus (HSV) associated with skin infections.
To evaluate the in vitro antimicrobial activity of aqueous and methanol extracts of Odina wodier bark (OWB), a folk medicine, against representative bacteria, fungi and herpes simplex virus (HSV) associated with skin infections.
The OWB extract(s) was found to inhibit the isolates of Staphylococcus aureus, Bacillus subtilis, Pseudomonas aeruginosa, Klebsiella pneumonia, Escherichia coli at an MIC of 256–5000 μg ml−1 and Candida albicans at and above 4000 μg ml−1 by agar and broth dilution assays. The growth curve of Staph. aureus revealed the highest activity within 2–6 h of methanol extract (ME) exposure. Interestingly, the MTT and plaque reduction assay showed that the extracts can inhibit HSV-1 and HSV-2 at EC50 of 22·4 and 28·8 μg ml−1, with Selectivity index of 11·7–15. While the time kinetic and binding assays demonstrated that the ME at 50 μg ml−1 prevents viral attachment into Vero cells. Phytochemical and HPLC analysis of ME revealed the presence of flavonoids, phytosterols, saponins and tannins including the pseudotannin chlorogenic acid.
The traditional use of OWB for the management of skin infections has scientific basis.
This study demonstrated the antimicrobial potential of OWB on selected isolates of bacteria, fungi and HSV, associated with skin infections.
The skin is the largest organ of human body accounting to 15% of total body weight that serve as an environmental interface and a protective covering. As surface tissues skin and mucous membranes are constantly in contact with outer environment and readily colonized by various bacteria, fungi and viruses. The skin and mucosal infections are common (Desta 1993) due to lack of sanitation, potable water, proper food and hygiene. Being the first line of defence against injury, skin damage leads to the invasion of a variety of pathogenic microbes (Robert and Kupper 1999) that account for 34% of all occupational diseases (Stevenson 1989; Spiewak 2001), prevalent in the working environment. For example, skin infection with haemolytic streptococcus is common in the slaughterhouse, parapox virus in Cows cause milker's nodules in milkman (Stevenson 1989) and sex workers suffer from HIV and its coinfection with HSV (Todd et al. 2013). The skin and mucosal infections caused by Staphylococci and Streptococci include wound, furuncles, carbuncles, abscesses, impetigo and erysipelas, while Pseudomonas aeruginosa prevalently cause wound and burn infections (Lory 1990; Murray et al. 1990). Sometimes E. coli may cause wound infection, but Candida albicans and Candida krusei are opportunistic pathogens to cause impetigo and candidacies (Madigan et al. 2003). However, HSV-1 and HSV-2 caused chronic and recurrent orofacial, and genital herpes that silently transmitted through close personal contact (Sucato et al. 1998; Chattopadhyay and Khan 2008).
The search for newer antimicrobials is a global challenge, as microbes are becoming resistant to the available antimicrobics (Latha and Kannabiran 2006) as drug resistance is a natural phenomenon and microbes may develop resistance even without any exposure to a drug (Olofsson and Cars 2007; Galán et al. 2013). One way to tackle the problem of drug resistance is by developing new compounds with different mechanism of action, either from natural sources or by combinatorial chemistry. Search for alternative agents from natural source provides flexibility and diversity that may help in reducing the side effects and its use in combination as synergistic (Keith et al. 2005; Kong et al. 2008) or adjunct therapy may delay the development of resistance (Koehn and Carter 2005; Beghyn et al. 2008; Hunter 2008). Problem of drug resistance, environmental degradation, pollution and toxicity of orthodox medicines have necessitated exploring nature as the source of effective agents in the management of human infections (Chah et al. 2006; Chattopadhyay and Khan 2008). Moreover, till date there are only a handful of antiviral agents are available, which are costly, yielded drug-resistant mutants and have toxicity. The antiherpes drug acyclovir and the related analogs frequently yielded drug-resistant viruses (Kleymann 2003; Miserocchi et al. 2007) and adverse drug reactions in pregnancy (Narayana 2008), neonates and children (Sawyer et al. 1988). Therefore, there is an unmet need for cheap, readily available natural agents to prevent microbial infections, with minimum side effects.
Plants produce a diverse range of bioactive secondary metabolites with unmatched chemical diversity (Parekh and Chanda 2007) and medicinal properties (Bentley 1997; Savithramma et al. 2011). Tribal healers in different parts of India use herbal medicines to treat skin ailments including cuts, wounds, swelling, eczema and infections (Samy et al. 1998; Ghosh et al. 2012; Mukherjee et al. 2013). Earlier we have reported some interesting antimicrobial leads from ethnomedicines of different tribes of India (Ghosh et al. 2012; Mukherjee et al. 2013). The ethnomedicinal plant Odina wodier (OW) Roxb. (Anacardiaceae) is a tall tree of deciduous forests in India, and neighbouring countries, popularly known as Rhus olina in English (Chidanbarathanu 2006). The plant is used for curing several ailments since the ages of Ayurveda, as its bark is used in curing ulcer, heart diseases, skin infection and wound healing (Kirtikar and Basu 1999; Ushakumari et al. 2012) and bark powder as toothpowder (Chidanbarathanu 2006). The leaves are helpful in elephantiasis (Kirtikar and Basu 1999) and abnormal vaginal discharge (Muthul et al. 2006), while gum odina serves as a tablet binder and emulsifying agent (Dinda et al. 2012). A recent study with ethanol extracts of five plants including O. wodier revealed antifungal activity against Candida species (Prabhakar et al. 2008). Moreover, the essential oil of Schinus molle L. (Anacardiaceae) has potent activity on Staphylococcus aureus, E. coli and Ps. aeruginosa (Rocha et al. 2012). However, till date OW has not been scientifically validated for its antimicrobial potential against the microbes that infect skin. Thus, the present study aimed to evaluate the antimicrobial spectrum of OW bark extracts against some selected pathogenic microbes responsible for skin infections, with a qualitative and quantitative phytochemical screening of the active extract.
The stem bark of OW was collected from Paschim Medinipur, West Bengal, India, and was identified by Dr. R. P. Nandi, Director, Cinchona and other Medicinal Plant Research (CMPR), Mungpoo, Darjeeling, India, and the voucher specimen was deposited with the CMRP and the host institute. The collected samples were washed, shed-dried and powdered, and the coarsely powdered bark (200 g) was separately extracted with 95% methanol and water for 72 h at room temperature (Chattopadhyay et al. 2001), filtered and solvent-evaporated under reduced pressure in a Eyela Rotary Evaporator (Tokyo Rikakikai Co., Ltd, Tokyo, Japan) at 40–45°C and then dried as powder. The percentage yields (w/w) of powdered methanol and aqueous extract were 9·34 ± 0·46 and 14·76 ± 0·21%, respectively. A weighted amount of the dried extract(s) was dissolved in 0·1% (w/v) dimethyl sulfoxide (DMSO) and diluted in sterile distilled water for microbiological evaluation.
The crude extracts were subjected to phytochemical group tests including tannin (with 10% potassium dichromate or lead acetate or 5% ferric chloride), reducing sugar (Benedict's and Fehling's tests), steroids (Liebermann–Burchard test), terpenoids (Salkowski test), flavonoids (extract was hydrolysed with 10% sulfuric acid, extracted with diethyl ether and divided into three parts to test with sodium carbonate, sodium hydroxide and ammonium solution) and others, following standard methods (Chattopadhyay et al. 2002). Moreover, the presence of pseudotannin including chlorogenic acid was tested by treating 2 ml of diluted extract(s) with 2–3 drops of ammonia solution (10%), and the mixture was heated over a flame and exposed to air to develop a green colour an indication of the presence of chlorogenic acid (Evans and Trease 1985). The physicochemical characters (total ash, acid insoluble ash and water content) along with the behaviour of powdered sample, dissolved in different chemicals and exposed to visible and UV (312 nm) light, were studied for constant quality and better yield (Exarchou et al. 2000).
HPLC analysis of the bioactive methanol extract (ME) of OW bark was carried out by a Shimadzu liquid chromatography, consisting of binary LC-20AD pumps coupled with a SPD-M20A photo-diode array detector, and a Rheodyne 7725 injector equipped with a 20-μl loop (Olszewska 2007), and a reverse-phase C18 (250 × 4·6 mm, 5 μm), Zorbax column (Agilent Technologies, Santa Clara, CA, USA) at ambient (25°C) conditions. The mobile phase was 1% acetic acid in HPLC-grade water/methanol (30 : 70) with the flow rate of 1 ml min−1. The absorbance was monitored at 210 nm with the injection volume of 20 μl. The stock solutions of ME of OW bark and marker compound chlorogenic acid (CA; Sigma-Aldrich Co. LLC., St Louis, MO, USA) were prepared by dissolving 10 mg of extract or pure CA in 10 ml of methanol separately and filtered (0·45 μm membrane), and then 20 μl of the sample was injected into the HPLC system for analysis. Identification of CA was based on their retention time and comparison of their UV spectra, while quantitative analysis was achieved by five-point calibration curves at 5–1000 μg ml−1 using the equation Y = 6973·2X + 60941 for CA, where Y represents the area of the extract and X represents the concentration of CA. The regression coefficient value was 0·9955 for CA. Real samples were diluted accordingly to fit the dynamic linear range of the regression line when necessary, and all measurements were taken in triplicates.
Fully characterized human isolates of Staph. aureus, Ps. aeruginosa, Klebsiella pneumonia, Escherichia coli, Candida albicans, Candida tropicalis and Cryptococcus neoformans (Table S1) collected from the Department of Microbiology, Calcutta Medical Collage and Hospital; Department of Bacteriology, Calcutta School of Tropical Medicine; and the Department of Bacteriology, National Institute of Cholera and Enteric Disease, Kolkata. The quality control strains of Staph. aureus ATCC 25923 and E. coli ATCC 25922 were used.
Antimicrobial sensitivity were tested by the disc diffusion method (Chattopadhyay et al. 2001) in Mueller Hinton agar plates, containing an inoculum size of 106 CFU ml−1 for bacteria or 2 × 105 for fungal spores cultured on Saboraud glucose agar plates. Previously prepared extract (aqueous and methanol)-impregnated discs at concentrations of 0–5000 μg ml−1 for bacteria and 0–64 mg ml−1 for yeasts or fungi were placed aseptically on sensitivity plates with appropriate controls (Chattopadhyay et al. 1998, 2001). All the plates were then incubated at 37°C overnight for bacteria or at 30°C for 3 days for fungi. The sensitivity was recorded by measuring the clear zone of growth inhibition on agar surface around the discs.
Minimum inhibitory concentration (MIC) was determined by agar and broth dilution methods (Chattopadhyay et al. 1998). A twofold serial dilution (0–5000 μg ml−1 for bacteria and 0–64 mg ml−1 for fungi) of the extracts was prepared in Mueller Hinton broth (bacteria) or Saboraud glucose broth (fungi). For agar dilution assay, previously prepared sensitivity plates, using serial two-fold dilutions of the extracts, were spot-inoculated (2 × 106 CFU per spot for bacteria and 2 × 105 spores per spot for fungi). The inoculated plates were then incubated at 37°C for 24 h (bacteria) or 30°C for 96 h (fungi). While for broth dilution tests, 0·1 ml of standardized suspension of bacteria (106 CFU ml−1) or fungal spores (5 × 105 spores ml−1) was added to each tube containing extracts (final concentration of 0–5000 μg ml−1 for bacteria and 0–64 mg ml−1 for fungi) and incubated with shaking, either at 37°C for 24 h (bacteria) or at 30°C for 96 h (fungi). The lowest concentration of the plate or tube which did not show any visible growth after macroscopic evaluation was considered as the MIC (Chattopadhyay et al. 2001).
Minimal bactericidal concentration (MBC) was determined by broth dilution method (Chattopadhyay et al. 1998). Standardized suspension (1 ml) of bacteria (2 × 106 CFU ml−1) or fungal spores (4 × 105 spores ml−1) was added into either Mueller Hinton broth or Saboraud glucose broth containing the MEat a final concentration 0- to 4-fold MIC. The mixtures were then incubated either at 37°C for 18 h or at 30°C for 96 h with shaking at 200 rpm on a platform shaker. The aliquots (1·0 ml) were withdrawn at different time intervals for the determination of OD at 540 nm as well as for the colony count, while the growth of the fungi was determined by the dry weight basis, after drying the sample at 60°C for 20 h consecutively for 3 days (Chattopadhyay et al. 2001).
African green monkey kidney cells (Vero cells, ATCC, Manassas, VA, USA) was grown and maintained in Eagle's minimum essential medium (EMEM), supplemented with 5–10% foetal bovine serum (FBS). After plaque purification, the standard strains of HSV-1F and HSV-2G (ATCC) were grown. The virus stocks were stored at −80°C for future use, and whenever required the virus stock was grown on Vero cells to determine the titres and used for further study.
The effect of OW bark extracts on Vero cell morphology was determined by MTT assay. Vero cells were cultured onto 96-well plates at 1·0 × 105 cells per well, and different concentrations of the extracts were added to each well at a final volume of 100 μl, in triplicate using DMSO (0·1%) and acyclovir (0–50 μg ml−1) as a negative and positive control, respectively. The drug-treated cells were incubated at 37°C with 5% CO2 for 2 days, and then the MTT reagent (10 μl) was added to each well. After 4 h of incubation, the formazan was solubilized by adding diluted HCl (0·04 mol l−1) in isopropanol, and the absorbance was read at 570 nm with a reference wavelength of 690 nm by an ELISA reader. Data were calculated as the percentage of cell viability by the formula: [(sample absorbance−cell free sample blank)/mean media control absorbance)]/100%. The 50% cytotoxic concentration (CC50) causing visible morphological changes in 50% of Vero cells with respect to cell control was determined (Zhang et al. 2007; Bag et al. 2012; Mukherjee et al. 2013).
Vero cells seeded in 12-well plates (5 × 105 cells per well) were treated with serial dilutions of the both aqueous and methanol extracts for 15 min at 37°C and then challenged with HSV-1 and HSV-2 (100 PFU per well) for 1 h. The inocula and drugs were subsequently removed from the wells, and the cells were washed with PBS twice and then overlaid with medium containing different dilutions of the extracts. After further incubation for 72 h, the supernatant was removed; the wells were fixed with methanol and stained with Giemsa (Sigma-Aldrich Co. LLC). The viral inhibition (%) was calculated as: [1−(number of plaques) exp/(number of plaques) control] × 100%, where ‘(number of plaques) exp’ indicates the plaque counts from virus infection with test extracts treatment and ‘(number of plaques) control’ indicates the number of plaques derived from virus-infected cells with control (0·1% DMSO) treatment (Cheng et al. 2003). The 50% effective concentration (EC50) for antiviral activity was defined as the concentration of extract that produced 50% inhibition of the virus induced plaque formation (Chattopadhyay et al. 2009; Bag et al. 2012).
Vero cells seeded in 96-well plates were infected with HSV-1 and HSV-2 at MOI of 1·0 in presence or absence of both the extracts at various concentrations (0, 5, 10, 25 and 50 μg ml−1). MTT assay was carried out after 2 days to determine the inhibition of infection, as described previously (Cheng et al. 2004; Bag et al. 2012). Values were obtained from three independent experiments with each sample performed in triplicate.
The effect of extract treatment over time was assessed according to the method of Madan et al. (2007) with some modifications. Briefly, to assess the effect of extract pretreatment, Vero cell monolayers seeded in 96-well plates were treated with ME (50 μg ml−1) for 24 h (long term) or 1 h (short term), washed with PBS and then challenged with HSV-1 (MOI 1) in EMEM containing 2% FBS. To study the effect of ME and virus concurrently, Vero cells were treated simultaneously with HSV-1 (MOI 1) and ME. After incubation for 1 h at 37°C, the virus–extract mixture was removed, and cells were washed prior to overlay with the fresh media. To evaluate whether the ME had any effects on or after viral entry, Vero cells were challenged with HSV-1 (MOI 1) for 1 h, and after removal of the inoculums, infected cells were washed and overlaid with fresh media containing extract (EC100 concentration). For the continuous drug treatment, cells were pretreated for 1 h with the extract, challenged with HSV-1 in presence of the drugs and overlaid with media containing the test extract after viral entry. MTT assay was carried out after 2 days to determine the inhibition of infection, as described previously. DMSO (0·1%) treatment was included as control in each experiment, as the powdered ME was dissolved in 0·1% DMSO and then diluted in sterile distilled water (Lin et al. 2011; Mukherjee et al. 2013).
Herpes simplex virus-1 (104 PFU ml−1) was mixed with ME at 50 μg ml−1 and then incubated at 37°C for 1 h. The extract–virus mixture was then diluted 100-fold (final inoculum, 100 PFU per well) with DMEM containing 2% FBS to yield a subtherapeutic concentration of the extract, and the virus inocula were subsequently added to Vero cells monolayers seeded in 12-well plates. For comparison, HSV-1 mixed with ME was diluted immediately to 100-fold (no incubation period) and added to Vero cells for infection. The 100-fold dilution served to titrate the extract below their effective doses and prevent meaningful interactions with the host cell surface. After adsorption for 1 h at 37°C, the diluted inocula were discarded, and the cells were washed twice with PBS. Then, the wells were covered with an overlay medium (DMEM containing 2% FBS) and were incubated at 37°C for 72 h before being subjected to plaque assay (Lin et al. 2011; Mukherjee et al. 2013). Viral plaques were counted, and plaque numbers obtained from infections set in presence of extract were compared with the DMSO (0·1%) control.
Vero cell monolayers grown in 12-well plates were prechilled at 4°C for 1 h and subsequently incubated with HSV-1 (100 PFU per well) for 3 h at 4°C to allow viral adsorption. The infected cell monolayers were then incubated with ME (50 μg ml−1), heparin (100 g ml−1) or DMSO (0·1%) for an additional 20 min at 37°C to facilitate HSV-1 penetration. At the end of the incubation period, extracellular virus was inactivated by citrate buffer (pH 3·0) for 1 min, and then the cells were washed with PBS and overlaid with DMEM containing 2% FBS. After 48 h of incubation at 37°C, viral plaques were stained and counted (Madan et al. 2007; Mukherjee et al. 2013).
Results were expressed as mean SEM (n = 6), and the statistical analyses were performed with one-way analysis of variance (anova) followed by post hoc Dunnett's test. A value of P < 0·05 was considered to be statistically significant compared with the respective control.
The ME of the plant part (sample) collected in February had the minimum amount of water (24·69 ± 2·50%), total ash (4·55 ± 0·55%) and acid insoluble ash content (0·24 ± 0·33%) and gave maximum yield (12·10 ± 0·51%). Under visible and UV254 nm light, the colour of powdered extracts was reddish to light magenta. Preliminary phytochemical screening of both the extracts revealed the presence of tannins, flavonoids, phenols, phytosterols, glycosides, saponins and reducing sugar including pseudotannin. Moreover, the HPLC quantification revealed that the ME of OW bark contains about 0·33% (w/w) of chlorogenic acid (CA) and pseudotannin, and we used CA as marker compound because the tannin was found to be the major group in the extract. Till date there was no report of the presence of tannin or CA from this plant. The HPLC chromatograms of the extract and standard CA were presented in Fig. 1a,b.
Methanol and aqueous extract of the OW bark afforded, upon removal of the solvent, a dark viscous reddish powder which demonstrated good inhibitory activity against five drug-resistant isolates of bacteria and one fungus that infect skin. The antimicrobial activity of methanol and aqueous extract tested against 96 bacterial and 3 fungal strains showed significant inhibitory activity with MIC bellow 2000 μg ml−1 and 16 mg ml−1, respectively (Table 1). Altogether 18 strains of Staph. aureus, 9 of Bacillus sp, 9 of E. coli, 2 of Kl. pneumonia and 2 of Candida species were inhibited. However, four strains Staph. aureus was inhibited by the ME at 256 μg ml−1 (Table 1), while nine strains each of E. coli and Bacillus sp., two of Kl. pneumonia and 18 strains of Staph. aureus were inhibited by the aqueous extract at an MIC of <5000 μg ml−1 (Table 1). The results further revealed that the MIC of ME against C. albicans and C. tropicalis was 4 and 8 mg ml−1, respectively, whereas C. neoformans had an MIC of 32 mg ml−1 (Table 1). Hence, the results indicated that the OW bark extract(s) possesses moderate to poor degree (10–15 mm) of antifungal activity against two Candida species, while C. neoformans had weak activity (7–10 mm) at and above 4 mg ml−1. Moreover, our preliminary screening showed that the aqueous extract had relatively less antimicrobial activity compared with the ME.
|Bacteria||No of strains tested||MIC (μg ml−1)||Zone of inhibition (mm) at MIC concentration||Antibiotic resistance|
|ME extract||AQ extract||ME extract||AQ extract||Pattern|
|Staphylococcus aureusa||4||256||1000||17·3–13·5||12·2–8·9||C, OFX|
|Staphylococcus aureus a||8||512||2000||13·8–9·8||9·6–7·5||NOR, Q, CIP|
|Staphylococcus aureus||5||2000||4000||9·9–7·8||8·5–6·2||NA, ATM|
|Streptococcus faecalis||2||4000||–||7·5–6·8||–||All sensitive|
|Bacillus subtilis a||3||512||2000||15·2–10·4||12·7–10·2||All sensitive|
|Pseudomonas aeruginosa||1||3000||–||8·7||–||A, C, CIP|
|Escherichia coli a||2||1000||3000||10·3–8·8||7·4–6·3||A, AMX, T|
|Escherichia coli||6||2000||5000||8·2–7·4||6·8–5·9||ATM, C, Q|
|Escherichia coli||12||5000||–||6·8–5·4||–||CTX, Fx, NA|
|Klebsiella pneumonia||2||2000||4000||8·2–6·3||7·1–6·4||All sensitive|
|Staphylococcus aureus ATCC 25923a||1||256||1000||13·6||10·4||All sensitive|
|Escherichia coli ATCC 25922||1||2000||4000||8·2||7·8|
|Candida tropicalis||1||8000||32 000||10·6||7·2||Sensitive|
|Cryptococcus neoformans||1||32 000||64 000||8·2||5·6||Sensitive|
Further, the disc diffusion assay showed that the different concentration of aqueous and ME of OW bark had different zone of inhibition at their MIC against the tested bacterial and fungal isolate (Table 1). The results presented in Table 1 also revealed that both the extracts exhibit highest zone (12·2 and 17·3 mm) of inhibition against Staph. aureus isolate(s) (Fig. S1).
Minimal bactericidal concentration assay, using 0- to 4-fold MIC, revealed that the ME had bacteriostatic or fungi-static activity at lower concentrations but cidal at higher concentrations, probably due to the interference by one or more active principle(s) of the extract. The growth inhibitory study revealed that the growth of Staph. aureus ATCC 25923 was decreased by 3-fold concentration of MIC, while growth of the clinical isolate of Staph. aureus, Bacillus subtilis (Fig. 2), C. albicans and C. tropicalis (Fig. 3) was decreased at 4-fold concentrations of their MIC, indicating that their MBC was 3- to 4-fold higher than their MIC. Moreover, as a representative, the growth curve of Staph. aureus demonstrated that the growth of the bacterium was inhibited at its MIC, within 2–6 h of its exposure to the extract, compared with the control strain used (Fig. 4).
The cytotoxicity assay with aqueous and ME of OW bark indicated that the extracts had no cytotoxicity up to 300 μg ml−1 on Vero cells, while a dose-dependent cytotoxic effect was observed above 300 μg ml−1 of both the extract, and the 50% cytotoxicity (CC50) of aqueous and methanol extracts was 398·6 and 336·5 μg ml−1, respectively (Table 2).
|Test drug||CC50a||EC50b||Selectivity index (SI)c|
|Aqueous extract||398·6 ± 3·22||62·2 ± 2·1||74·6 ± 1·8||06·4||05·3|
|Methanol extract||336·5 ± 4·84||22·4 ± 3·6||28·8 ± 2·5||15·0||11·7|
|Acyclovir||128·8 ± 3·44||2·6 ± 0·1||2·90 ± 1·3||49·5||44·4|
We then evaluated the antiviral effects of both methanol and aqueous extracts against HSV-1-infected Vero cells (PFU 100) by plaque reduction assay, using acyclovir as positive and DMSO (0·1%) as negative control. The results revealed that both aqueous and methanol extracts could inhibit viral plaque formation in a dose-dependent manner, and the 50% effective concentration (EC50) was 62·2 and 22·4 μg ml−1 for HSV-1, and 74·6 and 28·8 μg ml−1 for HSV-2, respectively (Table 2). The selectivity index (SI), a measure of the preferential antiviral activity of a drug in relation to its cytotoxicity (CC50/EC50), of aqueous and methanol extracts was calculated to be 6·4 and 15·0 for HSV-1 and 5·3 and 11·7 for HSV-2 (Table 2). Thus, based on higher SI value, methanol extract was selected for subsequent study.
Dose–response assay was conducted to know the concentration at which the ME inhibits 100% growth of the virus (EC100). For this, Vero cells (1 × 105) were infected with HSV-1F and/or HSV-2G (moi: 1·0) in presence of various concentrations of the ME for 48 h. The results revealed that the extract at 50 and 60 μg ml−1 completely inhibits (>99%) both HSV-1 and HSV-2, respectively, in a dose-dependent manner (Fig. 5), whereas acyclovir inhibits HSV-1 at 5 μg ml−1 and HSV-2 at 10 μg ml−1. Here, 0·1% DMSO was used as negative control.
To understand the possible mode and mechanism of antiviral activity of the ME on HSV-1 infection, we conduct time of addition and removal assay of the extract at its EC100 dose (50 μg ml−1). The extract was added at different time points of the virus life cycle (pre-entry, entry, and postentry), and efficacy was determined by MTT assay. Results demonstrated that pretreatment of Vero cells with extract (both long and short term) unable to protect the cell against HSV-1 infection. However, the extract was effective in preventing cell distraction when added during virus adsorption, immediately after viral entry, and throughout multiple cycles of virus replication (Fig. 6). The data also indicated that HSV-1 infection is severely impaired when the extract was present at the time of infection or during viral spread. Thus, it is unlikely that the antiviral activity of ME is due to its direct effects on the cells by masking cellular receptors or entry factors of HSV-1.
To evaluate the antiviral mode or mechanism of action of ME, we investigated its effect on virus entry, particularly the attachment and penetration steps of HSV-1 life cycle. The results showed that when the extract was pre-incubated with the virion and then diluted to a subtherapeutic concentration before infection, it interacts with the virion irreversibly and prevented infection (Fig. 7a). This suggests that the extract can bind to virion and neutralize its infectivity.
To assess the effects of the extract on virus penetration, HSV-1 particles were allowed to bind with the Vero cells at 4°C, subsequently fuse and penetrate the host cell membrane by shifting the temperature to 37°C in the presence or absence of the extract. As shown in Fig. 7b, the extract retained most of its antiviral activity even during the viral penetration and could completely abrogate virus penetration into the Vero cells, resulting in a protected cell monolayer. In contrast, heparin is known to block HSV-1 adsorption (McClain and Fuller 1994), but unable to prevent subsequent penetration (Lin et al. 2011), which is consistent with our results. Thus, our results collectively suggest that the tested extract may bind to the virus particles and thereby inhibited the attachment and penetration of HSV-1 during infection.
In this study, we examined the in vitro antimicrobial activity of aqueous and methanol extracts of an ethnomedicinal plant OW bark on selected isolates of bacteria, fungi and HSV that have direct or indirect involvement in skin infections. The importance of this study lies on the ethnomedicinal use of OW bark against skin infection which is not yet validated and the endangered nature of the plant. The phytochemical study showed that the bioactive extract contains phytoconstituents including tannin, flavonoids, saponins, while the HPLC chromatogram of ME revealed the presence of tannin and pseudotannin including the chlorogenic acid as one of the major group. It is known that chlorogenic acid has antimicrobial activity (Wang et al. 2009; Lou et al. 2011), while some tannins have antiviral activity also (Lin et al. 2011). Moreover, tannin and related polyphenolic groups including flavonoids are soluble in water, but methanol can serve as a better solvent for chlorogenic acid, tannins and flavonoids, as evident from a recent study with the extracts of Hazelnut shell (Nazzaroa et al. 2012). Thus, our study for the first time demonstrated the antimicrobial activity of OW bark extract containing tannin, chlorogenic acid and related polyphenolic groups and provides evidence to support its efficacy, particularly on some pathogenic microbes that may cause skin infections. Due to the increasing level of antimicrobial resistance of pathogenic microbes, particularly the organisms that infect skin and mucous layer, the choice of effective and safe treatment is becoming limited. Thus, traditional or ethnomedicinal plant extract or their phytochemical(s) need to be evaluated for therapeutic use.
The results revealed that the ME had higher antimicrobial activity than the aqueous extract, probably due to the different solubility of the active component in different solvents (Karou et al. 2007). Moreover, different isolates of bacteria or fungi exhibited varying degree of sensitivity to the ME, which is consistent with the earlier finding of Anani et al. (2000) that extract of Sida acuta had a significant activity on Staph. aureus, E. coli, B. subtilis and Mycobacterium phlei, but not on Streptococcus faecalis and Kl. pneumoniae. This difference in susceptibility can be attributed to either inherent resistant factor of different isolates or the previous exposure of the organism to other antimicrobial agents due to drug abuse in the population. Further, it was found that the highest concentration of the extracts had moderate to poor effect on the fungal isolates, with lower zone of inhibition, compared with the control. This is similar to the observation of Penduka and Okoh (2012) that the MIC of Garcinia kola methanol extract was lower in most bacteria than those for fungi. This is probably due to the structural and biochemical differences in the cells or cell wall of the tested microbes. We observed that the MIC of our extract varies between 250 and 5000 μg ml−1, and even the MIC of the same extracts varied against the different strains, although some strains had the same susceptibility pattern. Thus, it can be postulated that the tested microbes have different intrinsic levels of tolerance to antimicrobials that may cause the variation in MIC values among the isolates with relatively similar susceptibility patterns, as reported earlier (Ahmad and Aqil 2007).
Till date there is no cure of most of the viral diseases, including the mucocutaneous infections caused by HSV. Thus, the development of new antiviral molecules capable of inhibiting herpes virus infection represents an attractive strategy, particularly in immunocompromised individuals who often generate ACV-resistant HSV strains. Our results showed that the ME of OW bark has better activity than aqueous extract, and it effectively inhibits HSV infection in Vero cells without reducing the host cell viability. The results of viral entry, attachment and penetration assay showed that viral attachment and/or penetration is inhibited only when the extract and HSV virion are in contact with each other. Pretreatment of host cells with the extract, followed by removal of unabsorbed extract, indicated that masking cell surface receptors or entry factors by the extract or its phytoconstituents such as tannins are unlikely, while viral binding assays revealed that the extract blocked viral attachment to the host cell. Thus, our extract may bind to one or more glycoproteins of the infectious virions and make them inert, leading to the impaired glycoprotein function and there by prevent attachment and entry of the virion to the host cell. It is known that, during HSV-1 entry into epithelial cells, HSV-1 glycoprotein's gB, gC, gD, gH and gL interact with host cell receptors heparin sulfate, HVEM, nectin-1 and nectin-2 (Spear 2004) in an ordered and concerted manner to penetrate the host cell membrane (Spear 2004; Reske et al. 2007). As the viral entry and spread require a particular combination of viral surface proteins, the HSV glycoproteins gB and gD are repeatedly involved in viral binding, fusion and cell-to-cell transmission (Spear 2004; Reske et al. 2007; Heldwein and Krummenacher 2008). The associations between viral glycoproteins that mediate HSV-1 attachment and entry represent a complex scenario, and the targets of our extract may involve those glycoproteins during viral adsorption and penetration. We observed that the extract blocked virus attachment to Vero cells, such as heparin, suggesting that interaction of gC and gB with heparin sulfate proteoglycans is targeted by the extract to prevent virus internalization into cells in postbinding phase (Fig. 7b). Finally, it may be possible that viral glycoproteins still be accessible to the extract, even when these proteins are bound to the host cell or expressed in the intercellular junctions (McClain and Fuller 1994). Thus, the use of OW bark extract may help to improve the prognosis of anti-HSV therapy and reduced the risk of drug resistance by lowering its dose. However, further studies to determine the effectiveness of this natural product against other microbes and members of the herpesvirus family that infect skin are required.
Our study for the first time suggests that the traditional medicament OW bark may offer some beneficial effects in the management of skin infections caused by the tested bacteria, fungi and HSV, probably due to the interaction of one or more groups of phytoconstituents such as triterpene, flavonoids, tannins including chlorogenic acid, as detected in the HPLC chromatogram of the active extract.
The authors deeply acknowledged the Officer-In Charge, ICMR Virus Unit and the Head, Department of Pharmaceutical Technology, Jadavpur University, for laboratory facility.
No conflict of interest declared.